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Journal of Experimental Botany, Vol. 66, No. 22 pp. 6957–6973, 2015
doi:10.1093/jxb/erv415 Advance Access publication 9 September 2015
DARWIN REVIEW
Plastid RNA polymerases: orchestration of enzymes with
different evolutionary origins controls chloroplast biogenesis
during the plant life cycle
Thomas Pfannschmidt1,2,3,4,*, Robert Blanvillain1,2,3,4, Livia Merendino1,2,3,4, Florence Courtois1,2,3,4,
Fabien Chevalier1,2,3,4, Monique Liebers1,2,3,4, Björn Grübler1,2,3,4, Elisabeth Hommel1,2,3,4 and Silva Lerbs-Mache1,2,3,4
1 Université Grenoble-Alpes, F-38000 Grenoble, France
CNRS, UMR5168, F-38054 Grenoble, France
3 CEA, iRTSV, Laboratoire de Physiologie Cellulaire & Végétale, F-38054 Grenoble, France
4 INRA, USC1359, F-38054 Grenoble, France
2 * To whom correspondence should be addressed. Email: [email protected]
Received 9 July 2015; Revised 9 August 2015; Accepted 11 August 2015
Editor: Christine Foyer
Abstract
Chloroplasts are the sunlight-collecting organelles of photosynthetic eukaryotes that energetically drive the biosphere of our planet. They are the base for all major food webs by providing essential photosynthates to all heterotrophic organisms including humans. Recent research has focused largely on an understanding of the function of
these organelles, but knowledge about the biogenesis of chloroplasts is rather limited. It is known that chloroplasts
develop from undifferentiated precursor plastids, the proplastids, in meristematic cells. This review focuses on the
activation and action of plastid RNA polymerases, which play a key role in the development of new chloroplasts from
proplastids. Evolutionarily, plastids emerged from the endosymbiosis of a cyanobacterium-like ancestor into a heterotrophic eukaryote. As an evolutionary remnant of this process, they possess their own genome, which is expressed
by two types of plastid RNA polymerase, phage-type and prokaryotic-type RNA polymerase. The protein subunits of
these polymerases are encoded in both the nuclear and plastid genomes. Their activation and action therefore require
a highly sophisticated regulation that controls and coordinates the expression of the components encoded in the
plastid and nucleus. Stoichiometric expression and correct assembly of RNA polymerase complexes is achieved by
a combination of developmental and environmentally induced programmes. This review highlights the current knowledge about the functional coordination between the different types of plastid RNA polymerases and provides working
models of their sequential expression and function for future investigations.
Key words: Chloroplast biogenesis, developmental regulation, gene expression, nucleo–plastid interaction, plants, plastid RNA
polymerases.
Introduction
Today’s plastids are a group of morphologically and functionally diverse cellular organelles that are surrounded by a double membrane and are found specifically in plant and algae
cells. The most prominent representative of these organelles
is the chloroplast, which is found in green tissues of vascular
plants, mosses, and green algae. This plastid type is the site of
photosynthesis, but also of nitrate and sulphate fixation, and
parts of many other important cellular biosynthetic pathways
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6958 | Pfannschmidt et al.
including amino acid and lipid biosynthesis. Besides chloroplasts, other non-green plastid forms exist in plants such as
amyloplasts and elaioplasts, which are important for starch
and oil accumulation, respectively, in roots and storage
organs, or chromoplasts, which generate carotenoids responsible for the colours in fruits or flowers. All these different
plastid types develop from a common undifferentiated precursor form, the proplastid, which is also the form involved
in cytoplasmic inheritance. The respective plastid type that
is generated from this proplastid is determined by the tissue
context of the cell in which it resides. Furthermore, the different plastid types are convertible into each other, even after
differentiation, when respective environmental and/or cellular influences change (e.g. the chloroplast-to-chromoplast
conversion upon fruit ripening or in the opposite direction
following illumination). Thus, plant plastids display a remarkable morphological and functional diversity and flexibility
(Lopez-Juez and Pyke, 2005; Jarvis and Lopez-Juez, 2013).
Plant plastids evolved from a monophyletic evolutionary
event called primary endosymbiosis whereby a heterotrophic
mitochondriated protist engulfed a photosynthetically active,
free-living cyanobacterium. In the course of evolution, this
cyanobacterium was ‘tamed’ by the host cell and transformed into a stable and inheritable cell compartment, the
plastid (Cavalier-Smith, 1999; Reyes-Prieto et al., 2007). This
included a massive horizontal gene transfer from the endosymbiont towards the genome of the host, reshaping its biochemical properties (Martin et al., 2002; Timmis et al., 2004;
Reyes-Prieto et al., 2006). Current genome analyses strongly
suggest that the process of endosymbiosis was facilitated
by an infection of the host cell with Chlamydiae (Ball et al.,
2013). The result of this complex evolutionary event was the
generation of the first photosynthetic eukaryote. This ancient
lineage split into three branches, the ‘green’ lineage (chloroplastida), the ‘red’ lineage (rhodophyta), and the glaucophyta. From the green lineage around 450 million years ago,
the green algae and plants developed. The benefit provided
by plastids was such an advantage that several secondary and
even tertiary endosymbiotic events occurred within the green
and red algae resulting in the generation of further ‘plastidiated’ eukaryotes such as the dinoflagellates, the diatoms, the
euglenoids, and the apicomplexa (Delwiche, 1999; Stoebe and
Maier, 2002). This fascinating complexity is one of the prime
examples of the power of evolution within the tree of life.
As a remnant of their cyanobacterial origin, today’s plastids still contain their own genome, the plastome, and a fully
functional machinery for its expression. The plastome is comprised of around 120 genes in plants and up to around 200
genes in some red algae; it encodes mainly components of the
photosynthetic and gene expression machineries (Sugiura,
1992; Martin et al., 2002; Green, 2011). This is, however, only
a small part of the original coding capacity of the genome
of the ancient cyanobacterial precursor. Recent proteome
analyses identified up to 1660 non-redundant proteins within
Arabidopsis thaliana plastids [available in the online databases
AT_CHLORO and the Plant Proteome Data Base (PPDB)],
indicating that the vast majority of the chloroplast proteome
is encoded in the nucleus, translated in the cytosol, and
subsequently imported into the organelle (Sun et al., 2009;
Ferro et al., 2010). The present protein catalogues are probably not complete, as bioinformatic predictions of transit peptides identify many more potential plastid-localized proteins
in the genome. Estimates range from 2000 to 3500 proteins
that may be present in plastids. Low abundance and spatiotemporal expression variations probably represent the major
constraints in detection and identification of the lacking proteins (Abdallah et al., 2000; Van Wijk and Baginsky, 2011).
The dependency of plastids on the import of proteins from
the cytosol gained the host cell full control over development
and differentiation of the organelle (i.e. the cell type can fully
determine the plastid type). Therefore, usually only one type of
plastid is found in a given cell. Rare exceptions are some cells at
an early developmental stage in variegation mutants that possess
both normal chloroplasts and aberrant plastids. Segregation of
these plastids during further cell division subsequently leads to
the formation of white and green patches or stripes in the leaves
of the mutants (Wetzel et al., 1994; Sakamoto, 2003).
On the other hand, all major plastid protein complexes
involved in photosynthesis or gene expression contain several
plastid-encoded subunits that are essential for the assembly of
these complexes (Allen et al., 2011; Pfalz and Pfannschmidt,
2013). Thus, the plastids retained some influence on the
build-up of their own protein machineries and are, therefore,
regarded as genetically semi-autonomous. The plastid protein
complexes, in consequence, represent an evolutionary patchwork of subunits encoded in two different genetic compartments. Stoichiometric production and correct assembly of
these subunits thus require a faithful and tight communication between the two compartments (Rodermel, 2001; Pogson
et al., 2008; Woodson and Chory, 2008). This communication
must coordinate the expression of nuclear genes for plastid
proteins (typically present in one or few gene copies in the
nucleus) with that of plastid-encoded genes (present in up
to around 100 plastome copies per plastid and up to ~100
plastids per cell). This huge difference in gene copy number
needs to be adjusted by the gene expression machineries in
the nucleus and plastids (Lopez-Juez and Pyke, 2005; Dietzel
et al., 2008). The RNA polymerases found within the different genetic compartments are among the key regulators of
this complex interplay. By selective transcription of genes
essential for the corresponding developmental stage of the
tissue, they initiate a coordinated adjustment of gene expression events according to the needs of the organelles.
The RNA polymerases and their associated regulatory proteins comprise factors of eukaryotic, prokaryotic, and phage
ancestry, and reflect the complex evolutionary integration of
a prokaryotic gene expression system into a eukaryotic host
cell. Most of our current knowledge about this interplay was
obtained by studying chloroplast biogenesis, while much less
is known about these processes in other plastid types (e.g. in
chromoplasts or amyloplasts), although some studies exist
(Kahlau and Bock, 2008; Valkov et al., 2009). For reasons of
simplicity, this review focuses largely on processes occurring
during the light-dependent transition from the proplastid/
eoplast to the chloroplast, accepting that some aspects of the
described events might be different in other plastid types.
Plastid RNA polymerases govern chloroplast biogenesis | 6959
The different types of plastid RNA
polymerase
Plastids possess two types of RNA polymerase: (i) one multisubunit, plastid-encoded prokaryotic-type RNA polymerase
(PEP), and (ii) two different single-subunit, nuclear-encoded
phage-type RNA polymerase(s) (NEP) (Shiina et al., 2005;
Liere et al., 2011; Börner et al., 2015). The first requires
interaction with sigma-like factors that mediate promoter
recognition as well as with additional PEP-associated proteins (PAPs) of various functions, all encoded by the nuclear
genome (Fig. 1) (Schweer et al., 2010; Lerbs-Mache, 2011;
Pfalz and Pfannschmidt, 2013; Yu et al., 2014; Chi et al.,
2015). A proper function of the plastid transcription machinery therefore requires first the action of the eukaryotic nuclear
RNA polymerase II, which is responsible for the expression
of the nuclear-encoded components. The development of
functional plastids, then, depends on the coordinated action
of the two different types of plastid RNA polymerases,
which initiate a sequential and highly organized expression
cascade of essential plastid genes followed by further maturation of transcripts and appropriate translation. The RNA
metabolism and transcript maturation as well as the regulation of plastid translation are highly sophisticated processes,
especially in mature chloroplasts, and involve a high number
of regulatory proteins (Barkan and Goldschmidt-Clermont
2000; Monde et al., 2000; Eberhard et al., 2002), which cannot be covered by this review. It is, however, important to keep
in mind that these levels of gene expression are tightly interlinked. While plastid RNA polymerases provide transcripts
for proteins (including transcripts for components of the
large and small ribosome subunits) and tRNAs as substrate
for translation (Williams-Carrier et al., 2014), plastid translation is, in turn, essential for the generation of plastomeencoded Rpo subunits. In addition, both levels are highly
affected by the number of plastid DNA copies available (Udy
et al., 2012). Transcription, RNA maturation, and translation thus exhibit a mutual interdependency that needs to be
highly coordinated. This coordination during differentiation
and build-up phases of the organelle is mediated primarily by
developmental programmes yielding the typical tissue-specific
plastid types such as chromoplasts, amyloplasts, or chloroplasts. In fully matured chloroplasts, a functional regulation
that depends on environmental cues such as light variations
becomes more dominant. This functional regulation is mediated mainly by the chloroplast itself, for instance by redox
signals from photosynthesis (Pfannschmidt et al., 2003). The
orchestration of the different transcription machineries, thus,
Fig. 1. Schematic overview summarizing gene location, protein trafficking, and transcriptional activity of all components of the plastid transcriptional
machinery and their mutual interdependency. The three genetic compartments of plant cells are depicted. Coloured boxes within them represent genes,
with ovals and circles corresponding protein components. The nomenclature of genes and proteins given in these symbols follows that given in the text.
Black arrows indicate the traffic of information from a gene via transcripts and translation (indicated by white boxes) towards the place of action of the
corresponding protein. Blue symbols: Nuclear genes of NEP enzymes are represented by boxes. Plastid-located NEPs (ovals) are depicted above their
target genes in plastids (classes II and III, including rpoA, -B, -C1, and -C2). Rpo subunits are translated within the plastid and generate the PEP core
complex. Green symbols: PAP genes in the nucleus are probably expressed as a regulon requiring an as-yet-unknown master regulator (orange oval) for
coordinated expression. PAP subunits assemble around the PEP core. Relative positions of PAPs are based on literature data describing protein–protein
interactions in a yeast two-hybrid system (see main text). PAPs drawn with a red line display either a true (full line) or potential (broken line) second
location in the nucleus. The still-unknown pathways of nuclear trafficking via a cytosolic or a plastid route are indicated by broken red arrows. Yellow
symbols: Nuclear-encoded plastid sigma factors are transported into the plastid and mediate specific promoter recognition for the PEP complex (blue
Rpo subunits assembled with all green PAPs) for the target genes (yellow boxes: classes I and II as well as sigma factor-specific genes psbN, atpH, and
ndhF). For definition of classes I, II and III see text.
6960 | Pfannschmidt et al.
requires the integration of developmental as well as physiological regulation networks and integrates signals from the
nucleus towards the plastid (anterograde signalling) and signals from the plastid towards the nucleus (retrograde or plastidial signalling) (Pogson and Albrecht, 2011).
The bacteria-like DNA-dependent RNA polymerase of
plastids (PEP)
Core complex
Transcriptional activity in plastids was already reported in the
1960s (Kirk, 1964), but the nature of this activity remained
obscure until molecular biology techniques allowed the
sequencing of plastid genes (Hudson et al., 1988; Igloi et al.,
1990). Nowadays, it is common knowledge that plastids possess a RNA polymerase complex of prokaryotic origin that is
composed of subunits homologous to the well-studied subunits of RNA polymerase from Escherichia coli or other bacteria. So far, all sequenced chloroplast genomes of vascular
plants have revealed the presence of genes that exhibit significant sequence homologies to bacterial RNA polymerase (rpo)
genes (Fig. 1) (Igloi and Kossel, 1992). A comparison was performed of structural Rpo homology between Arabidopsis and
E. coli at the protein level using Phyre2 and found homologies that ranged from 37% (RpoC2) to 85% (RpoA) and 95%
(RpoB, RpoC1) (with respect to the lengths of the proteins).
The genes for these proteins are typically arranged as a large
operon comprising the genes rpoB–rpoC1–rpoC2 and a single
gene rpoA being part of another operon that contains several
genes for ribosomal components (Green, 2011). These rpo
genes encode the catalytic subunit RpoB (or β-subunit), the
subunit RpoC1 (or β′-subunit), which has no clearly assigned
function yet, and the subunit RpoC2 (or β″-subunit), which
is believed to possess a DNA binding function. Most likely,
the genes rpoC1 and rpoC2 emerged from a split of the
original rpoC gene of the cyanobacterial ancestor. In addition, in dicotyledonous plants, an intron can be found within
the rpoC1 gene that is not present in monocotyledons. This
observation might suggest different evolutionary constraints
for the evolving transcription machinery in these two plant
clades (Igloi and Kossel, 1992).Whether this is connected
to specific functional differences of the RNA polymerase
subunits between the two clades and/or whether this reflects
an adaptation to the different developmental programmes
underlying chloroplast biogenesis in monocotyledons and
dicotyledons still needs to be elucidated. Interestingly, the
rpo genes in the green algae Chlamydomonas reinhardtii are
monocistronic units dispersed over the plastome, indicating
that the plastome organization within the green lineage was
not thoroughly maintained (Maul et al., 2002). The subunit
RpoA (or α-subunit) probably serves as a stabilizing subunit
of the complex and, in analogy to its bacterial counterpart,
is assumed to occur as a dimer, although direct supporting
evidence does not exist. The basic RNA polymerase complex with the stoichiometry α2, β, β′, β″ is referred to as the
core enzyme and is capable of transcriptional elongation in
vitro and probably also in vivo. Plastid knockout mutants in
tobacco or nuclear knockdown mutants in Arabidopsis, both
with structural or functional deficiency for any of these rpo
genes, display an albino or yellowish phenotype with arrested
plastid development, indicating the absolute requirement of
Rpo subunits for proper chloroplast development. All these
mutant plants have been reported to be viable on sucrose-supplemented medium, indicating that the missing chloroplast
function can largely be rescued by an external carbon source
(Allison et al., 1996; Hajdukiewicz et al., 1997; De SantisMaciossek et al., 1999; Chateigner-Boutin et al., 2008, 2011;
Zhou et al., 2009).
Sigma factors
The typical promoters of PEP-transcribed genes resemble
prokaryotic promoters and correspondingly comprise –35
(TTGACA)- and –10 (TATAAT)-like-sequence motifs
(Shiina et al., 2005; Liere et al., 2011; Chi et al., 2015). For
specific promoter recognition and transcription initiation, the
PEP core complex is thus dependent on the interaction with
prokaryotic-like sigma factors (Allison, 2000), which were
first identified by biochemical means (Bülow and Link, 1988;
Lerbs et al., 1988). In contrast to the other Rpo subunits, the
genes for these factors were found to be encoded in the nucleus
(Liu and Troxler, 1996; Tanaka et al., 1996). Like many
other genes that were originally located in the genome of the
engulfed cyanobacterium, they went through a horizontal
gene transfer towards the nucleus of the host cell and are
therefore under the control of the nuclear transcription
system. In Arabidopsis, six different sigma factors are known
and are designated Sig1–Sig6 (Fig. 1) (Schweer et al., 2010;
Lerbs-Mache, 2011; Chi et al., 2015). In contrast to mutants
with defects in the core components, single sigma-factor
T-DNA inactivation mutants in Arabidopsis do not display
severe phenotypic defects, pointing to a certain level of
functional redundancy between the factors. Exceptions to
this are sig2 and sig6 knockout mutants, which exhibit lightgreen phenotypes indicating a more pronounced role of the
encoded Sig2 and Sig6 factors in plastid transcription in the
early stages of chloroplast differentiation. Apparently, their
loss can only partly be replaced by the other sigma factors
during the early steps of seedling development. Studies from
several laboratories indeed have revealed partial redundancy
for sigma factor functions but also identified gene-specific
functions for these regulatory factors (Schweer et al., 2010;
Lerbs-Mache, 2011; Malik Ghulam et al., 2012). The specific
functions, gene expression profiling, regulation of sigma
activity/specificity by post-transcriptional modifications,
and some evolutionary aspects have already been described
and discussed in detail in a number of previous reviews
(Lysenko and Kusnetsov, 2005; Schweer et al., 2010; LerbsMache, 2011; Chi et al., 2015). This review will focus mainly
on some evolutionary aspects of sigma diversification and
functional integration into the eukaryotic host during land
plant evolution.
Evolutionary origin of plant sigma factors Previous
phylogenetic analysis of sigma factor genes indicated that,
in plants, they all derived from the principal cyanobacterial
sigma factor SigA (Hakimi et al., 2000; Mache et al., 2002;
Plastid RNA polymerases govern chloroplast biogenesis | 6961
Lysenko and Kusnetsov, 2005). This suggests that, out of the
multiple sigma factors present in the ancient cyanobacterium,
only the primary sigma factor SigA has been retained in
the chloroplast ancestor. Comparison of intron positions
of plant sigma factor genes further suggests that Sig1 and
Sig2 originated from this SigA common ancestor by gene
duplication and that all other sigma factors were later
derived from Sig2 (Lysenko and Kusnetsov, 2005). The early
evolutionary origin of Sig1 and Sig2 was recently confirmed
by showing the presence of the corresponding sigma factor
genes in the nuclear genomes of the liverwort Marchantia
polymorpha (Kanazawa et al., 2013; Ueda et al., 2013), of the
moss Physcomitrella patens (Hara et al., 2001; Ichikawa et al.,
2004), and of the spikemoss Selaginella moellendorffii (Banks
et al., 2011). In addition to these two genes, the nuclear
genome of M. polymorpha encodes two other different sigma
factors. One was named Mp_SIGX because it does not relate
to anyone of the six higher plant sigma factors (Kanazawa
et al., 2013). The other one found to be orthologous to Sig5
of higher plants. Thus, Sig5 has evolved over the same time
scale as Sig1 and Sig2 but probably does not originate from
Sig2, nor from the cyanobacterial ancestor, because none
of the cyanobacterial sigma factors group together with
Sig5 (Mache et al., 2002). Sig5 has the weakest similarity
to other plant sigma factors. Noticeably, A. thaliana Sig5 is
orthologous to E. coli sigma factor RpoH, which is known
to be implicated in heat-shock and stress responses of the
bacterium (Mache et al., 2002). It seems that this function has
been evolutionary conserved in basal and higher land plants,
as suggested by the induction of Sig5 gene expression under
several stress conditions in liverwort as well as in Arabidopsis
(Nagashima et al., 2004; Kanazawa et al., 2013). However,
the evolutionary origin of Sig5 remains to be elucidated.
Diversification of plant sigma factors during plant
evolution, and co-evolution of sigma factors and
promoters As mentioned above, results obtained so far
from non-vascular land plants reveal the presence of sigma
orthologues to higher plant factors Sig1, Sig2, and Sig5. The
other sigma factors, namely Sig3, Sig4, and Sig6, emerged
during theevolution of vascular plants after the separation of
lycophytes and euphyllophytes. They probably originated by
gene duplication from Sig2. Sig4 represents the most recent
acquisition of a new sigma factor. It has been found so far
only in Superrosidae (Rosales and Geraniales). Its absence
in currently known nuclear genomes of monocotyledonous
plants suggests that it evolved after the separation of dicots
and monocots.
The recent emergence of Sig4 is of special interest as it can
be related to a specific functional diversification of sigma function (Schweer et al., 2010; Lerbs-Mache, 2011; Chi et al., 2015).
While Sig2 and Sig6 seem to be more general factors required
for transcription initiation of many various genes or operons,
Sig3 and Sig4 exclusively recognize the psbN and atpH promoters (Sig3) (Zghidi et al., 2007) and the ndhF promoter (Sig4) in
Arabidopsis (Fig. 1) (Favory et al., 2005). The first insights into
functional specification of sigma factors in non-vascular land
plants came from a recent analysis of the plastid transcriptome
of a sig1 mutant of M. polymorpha. These results revealed
changes that are similar to those observed in mutants of higher
plants, suggesting conservation of principal sigma activity of
Sig1. However, the ndhF gene is also transcribed by Sig1 in liverwort. A comparison of ndhF promoter regions from liverwort
and higher plants reveals considerable alterations in promoter
elements between liverwort, several monocotyledons species,
and Arabidopsis, testifying to co-evolution of sigma factors
and promoters (Ueda et al., 2013). Co-evolution of Sig4 and
ndh gene expression is also testified by the concomitant lack of
Sig4 gene expression (Zhang et al., 2015) and the loss of ndh
genes from the plastid genome in some Geraniaceae (Blazier
et al., 2011; Weng et al., 2014).
Geraniaceae plastid genomes are highly rearranged (Weng
et al., 2014) and are characterized by remarkably elevated
evolutionary rates, especially in rpoB, rpoC1, and rpoC2 genes
(Guisinger et al., 2008; Weng et al., 2012). In addition, numerous gene duplications of Sig5 and Sig6 have been observed in
various species (Zhang et al., 2015). Therefore, Geraniaceae
represent an attractive system to study plastid–nuclear
genome co-evolution with respect to the PEP transcriptional
apparatus. Variations in genes for RNA polymerase subunits
have already been shown to be useful for taxonomy classification and phylogenetic analyses (Taillardat-Bisch et al., 2003;
Iyer et al., 2004; Lane and Darst, 2010). Using Geraniaceae
as a model system, coordinated evolution rates between interacting plastid and nuclear genes coding for PEP subunits and
sigma factors, respectively, have recently been shown (Zhang
et al., 2015). The authors suggested that this strong correlation of evolutionary rates might provide an explanation for
the observed plastome–genome incompatibility between species within the Geraniaceae.
Emergence of sigma factor phosphorylation as
a mechanism for regulating activity/specificity of
transcription Plant sigma factors are composed of
evolutionarily conserved C-terminal parts and variable
N-terminal parts. The N-terminal parts have probably been
acquired during the process of integration of the ancestral genes
after duplication into different regions of the host’s nuclear
genome. The N-terminal parts have functionally diverged
during evolution. They are incompatible with prokaryotic
transcription systems as shown for E. coli (Hakimi et al.,
2000; Mache et al., 2002), and they are not interchangeable
between different plant sigma factors (Schweer et al., 2009).
In contrast to their bacterial counterparts, plant sigma factors
can be modified post-translationally by phosphorylation, and
this modification seems to occur exclusively in the N-terminal
variable region. The functional consequences of sigma factor
phosphorylation have so far been investigated in vivo for
Sig1 (Shimizu et al., 2010) and Sig6 (Schweer et al., 2010)
in Arabidopsis; however, using bioinformatics, hypothetical
phosphorylation sites were predicted in all plant sigma
factors. Phosphorylation has been observed in Sig2 by an in
vitro approach, and phosphorylation of Sig1 and Sig6 have
been associated with changes in the binding efficiency of
RNA polymerase–sigma complexes to selected promoters
(Türkeri et al., 2012). Phosphorylation of Sig1 is regulated by
6962 | Pfannschmidt et al.
redox signals and serves to adapt photosystem stoichiometry
to daily changing light conditions via changes in psbA and
psaA transcript levels (Shimizu et al., 2010).
Direct phosphorylation of sigma factors represents a mechanism of transcriptional regulation that has been acquired
after the endosymbiotic event. However, it is not clear yet
whether this mechanism evolved before or after the separation of Viridiplantae and Rhodophyta. Western blot analyses
of the four sigma factors of the red alga Cyanidioschyzon
merolae (note that these sigma factors cluster together and
are not related to Sig1–Sig4 of higher plants as shown for
Cyanidium caldarium; Mache et al., 2002) indicate posttranslational modifications for Sig3 and Sig4 (Kanesaki et al.,
2012). The nature of these modifications is not known, but
phosphorylation might be considered.
PEP-associated proteins (PAPs)
While genetic data imply a prokaryotic structure of the plastid RNA polymerase complex, biochemical purifications of
this enzyme from chloroplasts yielded protein complexes with
a much higher subunit number (15–20 depending on the species). These multi-protein complexes thus resemble more the
eukaryotic-type RNA polymerases from the nucleus (Lerbs
et al., 1985; Rajasekhar et al., 1991). Biochemical studies on
mustard cotyledons then revealed that the enzyme in its simple
prokaryotic composition (i.e. α2, β, β′, β″) could be purified
from etioplasts of dark-grown seedlings. This complex can be
referred to as the PEP core complex and forms a holo-enzyme
together with the sigma factors for transcription initiation. It
is able to rapidly acquire additional subunits upon light exposure, which triggers chloroplast formation (Pfannschmidt and
Link, 1994). These additional subunits were subsequently
identified by mass spectrometry approaches in mustard,
tobacco, and Arabidopsis (Pfannschmidt et al., 2000; Suzuki
et al., 2004; Pfalz et al., 2006; Steiner et al., 2011). The corresponding genes were identified in the Arabidopsis nuclear
genome by homology searches and all displayed a predicted
chloroplastic transit peptide at the N terminus. The majority
of the subunits of the plastid-encoded RNA polymerase are
thus imported from the cytosol (see below).
A major challenge in studying the additional subunits of
the plastid RNA polymerase complex by biochemical means
is to discriminate between bona fide subunits from contaminating, irrelevant, or non-essential subunits. Plastid DNA and
the enzymes responsible for its replication and expression are
organized into huge structures resembling bacterial nucleoids.
These plastid nucleoids can be visualized in vivo by various
optical methods, including DAPI staining, revealing that they
are first located at the proplastid envelope and then attach
to thylakoids in fully differentiated chloroplasts (Krupinska
et al., 2014; Powikrowska et al., 2014). Although nucleoids
appear as distinctly shaped bodies in microscopy, they cannot
be isolated as such since they do not possess any enclosing
membrane or other stabilizing outer structure, and therefore
precise subunit catalogues are difficult to obtain. The best
approach so far is the purification of the transcriptionally
active chromosome (TAC), a huge DNA–protein complex
that is capable of in vitro RNA synthesis upon addition of
nucleotides. It consists of around 40–60 proteins and was analysed in detail by mass spectrometry in Arabidopsis, mustard,
and spinach (Pfalz et al., 2006; Melonek et al., 2012). A total
of 35 proteins were identified in Arabidopsis from which 18
were novel and were called pTACs (Pfalz et al., 2006). Many
of them are not involved in transcription directly but cover
other important functions including structural stabilization
of the DNA, membrane anchoring, RNA processing, and
ribosome formation. The plastid RNA polymerase complex
is an intrinsic part of this TAC, but it can also be purified
by alternative protocols as a soluble enzyme activity. This
enzyme complex is much smaller than the TAC and requires
exogenously added DNA as template for transcription (Pfalz
and Pfannschmidt, 2013). The smallest complex reproducibly purified from mustard chloroplasts comprises ten subunits in addition to the Rpo subunits (Steiner et al., 2011)
and was formerly assigned as peak A enzyme (Pfannschmidt
and Link,1994). The core complex consisting only of Rpo
subunits could be purified as peak B enzyme from etioplasts
or young immature chloroplasts (Pfannschmidt and Link,
1994). Complexes with intermediate subunit composition
have never been found, strongly suggesting that the large peak
A enzyme is only stable when all ten additional subunits are
present at the same time. It therefore can be defined as the
minimum RNA polymerase complex of chloroplasts, which
may be larger by transient addition of further subunits.
The potential functions of these ten subunits were predicted
by domain analysis of the corresponding genes in Arabidopsis
and verified in some cases by functional characterization of
the respective T-DNA inactivation mutants (see below for
detailed description). Intriguingly, all these mutants display
an albino/ivory or pale-green phenotype with arrested plastid development, no or minor PEP activity, and an enhanced
NEP activity. The mutations are seedling lethal on soil, but
the albino mutants can be maintained and grown on sucrosesupplemented medium (Schröter et al., 2010). These phenotypic defects are highly reminiscent of those of plants in
which PEP activity was inactivated by genetic approaches
(plastomic Δrpo mutants), indicating that the presence of
each single subunit is as essential for the activity of PEP as the
rpo-core subunits themselves. Therefore, it is postulated that
proteins that (i) are present in this minimum RNA polymerase complex, and (ii) cause severe defects in chloroplast biogenesis when inactivated (albinos) define a set of true RNA
polymerase subunits that have been named PEP-associated
proteins (PAPs) (Pfalz and Pfannschmidt, 2013). Not surprisingly these subunits were also found within the group of
pTAC proteins (Pfalz et al., 2006). This allows a distinction to
be made between true RNA polymerase subunits (PAPs) and
more peripheral components among the pTACs that are less
directly involved in transcription and for which chloroplast
biogenesis phenotypes in inactivation mutants are less pronounced or even wild-type like.
In Arabidopsis, so far 12 different PAPs have been identified that meet the mentioned criteria (Fig. 1). In co-expression analyses, they display a high degree of correlation in
their transcript accumulation patterns, suggesting that they
probably generate a regulon (Steiner et al., 2011). For PAP1,
Plastid RNA polymerases govern chloroplast biogenesis | 6963
it was demonstrated that both RNA and protein are rapidly
induced by illumination (Yagi et al., 2012). More detailed
in silico expression analyses were performed using available
databases at the eFP browser to test this observation for the
other pap genes. To this end, the average expression value for
the pap gene regulon under all stages of the life cycle (as far
as available) was calculated and compared it with the highest
expression value found in mature rosette leaves (set to 100%)
(Fig. 2, see bar diagram). In general, high pap transcript accumulation was found in green tissues containing chloroplasts
(such as carpel, sepals, rosette leaves, and the shoot apical
meristam. In contrast, only low expression was observed in
non-green tissues such as flower petals and stamen, pollen,
roots, and dry seeds (Fig. 2). Interestingly, imbibition of seeds
alone led already to significant induction of pap genes. This
suggests that pap genes might be induced not only by light
but also by internal developmental programmes. In summary,
the data indicated that pap genes are mainly expressed in
chloroplast-containing tissues and in tissues that are prone to
generate chloroplasts (Fig. 2). This coincides with the initial
biochemical observations describing a light-induced restructuring of the PEP complex upon etioplast-to-chloroplast
transition (Pfannschmidt and Link, 1994). It is thus conceivable that pap gene expression is enhanced by nuclear RNA
polymerase II as part of the developmental programmes
that serve seedling development and photomorphogenesis.
The controlling factors/regulators for this distinct response
remain to be determined but most likely are connected to
Fig. 2. Expression analysis of the PAP ‘regulon’ during the life cycle of Arabidopsis. Using the eFP browser (http://bar.utoronto.ca), the developmental
series of expression levels was obtained for an average of 11 PAPs (centre plot). Means and standard deviations were calculated and plotted as
percentages of the highest score set to 100% (marked by arrow head). The stage of life cycle tested can be identified by the positions numbered on the
x-axis. 1, Dry seed (ds); 2, imbibed seed, 24 h (imb); 3, first node (not shown on the life cycle); 4, flower stage 12, stamens (not shown); 5, cauline leaf (cl);
6, cotyledon (c); 7, root (r); 8, entire rosette after transition to flowering (not shown); 9, flower stage 9 (f9); 10, flower stage 10/11 (f10); 11, flower stage 12
(f12); 12, flower stage 15 (f15); 13, flower stage 12, carpels (not shown); 14, flower stage 12, petals (not shown); 15, flower stage 12, sepals (not shown);
16, flower stage 15, carpels (floral diagram); 17, flower stage 15, petals (floral diagram); 18, flower stage 15, sepals (floral diagram); 19, flower stage 15,
stamen (floral diagram); 20, flowers stage 15, pedicels (pl); 21, leaf 1 + 2 (not shown); 22, leaf 7, petiole (p); 23, leaf 7, distal half (d); 24, leaf 7, proximal
half (px); 25, hypocotyl (h); 26, root (r); 27, rosette leaf 2 (rl2); 28, rosette leaf 4 (rl4); 29, rosette leaf 6 (rl6); 30, rosette leaf 8 (rl8); 31, rosette leaf 10 (rl10);
32, rosette leaf 12 (rl12); 33, senescing leaf (sl); 34, shoot apex, inflorescence (ifm); 35, shoot apex, transition (ft); 36, shoot apex, vegetative (sam); 37,
stem, second internode (not shown); 38, mature pollen (mp); 39, seeds stage 3 with siliques (s3); 40, seeds stage 4 with siliques (s4); 41, seeds stage 5
with siliques (s5); 42, seeds stage 6 without siliques (s6); 43, seeds stage 7 without siliques (s7); 44, seeds stage 8 without siliques (s8); 45, seeds stage
9 without siliques (s9); 46, seeds stage 10 without siliques (s10); 47, vegetative rosette (arrowhead, reference set at 100%). Each value was then color
coded and attributed to the schematic features on the life cycle. F, fertilization; EG, embryogenesis (inset picture depicts embryos in their photosynthetic
phase); G, germination; PMG, photomorphogenesis (green arrow, light-dependent etioplast-to-chloroplast transition); FT, flowering transition; FMG; floral
morphogenesis; GG, gametogenesis; FG, female gametophyte. Colours range from yellow (no expression) to red (full expression as described above for
the plot); white, not documented. Displayed features were inspired by the eFP browser developmental map.
6964 | Pfannschmidt et al.
the processes in the early steps of germination and embryo
development.
Although all Arabidopsis pap knockout mutants display highly similar phenotypes, the predicted functions and
structures of the respective PAPs are quite diverse and bring
a multitude of functional as well as structural features to
the complex. For an exhaustive summary, all 12 pap gene
sequences including their predicted functional domains and
their corresponding predicted three-dimensional structure
around the E. coli RNA polymerase core complex have been
complied in Fig. 3 (for more details see the figure legend). Our
current working hypothesis is that all the PAPs are required
for the formation and/or stability of the PEP complex and
that the lack of any of these subunits is detrimental to the
whole complex, hence leading to defects broader than the one
corresponding solely to the functional loss of any particular subunit. According to structural prediction of functional
domains, the proteins can be subdivided into different groups
comprising gene expression- or redox-related functions and a
third group with unknown or metabolism-related functions.
In addition, our predictions identified numerous PAPs that
possess a nuclear localization signal (NLS) in addition to the
N-terminal chloroplastic transit peptide (PAP1, -5, -7, -8, -9,
and -12) (Fig. 3). They therefore probably belong to a special
group of proteins recently identified that are dually localized
to plastids and the nucleus (Krause and Krupinska, 2009).
However, only for PAP5/HMR/pTAC12 has a dual localization in the nucleus and plastids been shown so far, both in
Arabidopsis and in maize (Fig. 1) (Chen et al., 2010; Pfalz
et al., 2015). The functionality of the NLS in the other PAPs
awaits experimental proof. For the nucleus-localized PAP5/
HMR/pTAC12 of Arabidopsis, a function in photomorphogenesis has been attributed (Chen et al., 2010). The mutant
allele hmr-5 shows defects in photomorphogenesis corresponding to hypocotyl elongation under red light and an
albino phenotype that is typical of pap mutants (Chen and
Chory, 2011). It could be demonstrated that HMR physically interacts with phytochrome-interacting-factors (PIFs)
and mediates the degradation of PIF1 and PIF3 (Galvao
et al., 2012), known to be an important step in the transition from skotomorphogenesis to photomorphogenesis. A 9
aa transcriptional activation domain at the C terminus of
PAP5/HMR/pTAC12 mediates the expression of PIF target
genes (Qiu et al., 2015), indicating that PAP5/HMR/pTAC12
can directly affect gene expression in the nucleus. This coincides with the action of plastidial PAP5/HMR/pTAC12, as
shown recently in maize. It could be demonstrated that the
orthologous ZmpTAC12 possesses a binding activity for
Fig. 3. Predicted protein structure of the PAP proteins in Arabidopsis and a structural model of the PEP complex. PEP core: Structural model borrowed
from bacterial-type rpo core complex (Ebright lab at http://rutchem.rutgers.edu); PAP1–PAP12: yellow, chloroplastic transit peptide (cTP); red, bipartite
or monopartite NLS (bNLS or mNLS). PAP1: magenta, PNLR domain, stabilization of rbcL RNA; dark green, PPR (pentatricopeptide repeat) motif, RNA
binding involved in RNA maturation and translation; cyan, SAP domain, a putative DNA binding motif involved in chromosomal rearrangement. PAP2:
dark green, PPR domain composed of nine motives; light green, SMR small MutS related involved in mismatch repair. PAP3: cyan, S1-like domain,
involved in RNA binding. PAP4 and PAP9: cyan and sky blue, SOD Fe/Mn superoxide dismutase C-terminal and N-terminal domains, respectively. PAP5:
gold, glutamine-rich region; green, PEST signature of short-lived molecules (adapted from Chen et al., 2010). PAP6: green, fructokinase with substrate
and ATP binding site. PAP7: magenta, SET domain, involved in methyl transferase activity; cyan, Rubisco binding domain sharing similarity with histone
binding motives. PAP8: no homology with known functional domain. PAP10: magenta, thioredoxin domain. PAP11: pink and magenta, MurE ligase
domain, involved in the incorporation of amino acids into peptidoglycan. PAP12: no homology with known functional domains. Blue lines indicate protein
regions used for structural models predicted by Phyre2 (http://www.sbg.bio.ic.ac.uk/phyre2).
Plastid RNA polymerases govern chloroplast biogenesis | 6965
single-stranded nucleic acids, but without displaying a genespecific function (Pfalz et al., 2015). Its essentiality for PEP
complex formation could also be confirmed in this monocotyledonous plant. Interestingly, ZmpTAC12 exists in two
differentially migrating forms that are present in both the
nucleus and plastids. This poses questions for the trafficking
of these proteins within the cell (see below) and the impact
of the chloroplastidial PAP5/HMR/pTAC12 function on its
nuclear action; i.e. would a lack of the PEP complex interfere
with photomorphogenesis because the greening has become
impossible and how is the retrograde signalling impacted by
PAP5/HMR/pTAC12 loss of function?
Transcriptional expression of the PAPs has been observed
not only in green tissues but also in tissues preceding their
greening, e.g. during imbibition of seeds (Fig. 2), whereas the
fully assembled PEP complex has only been observed in lightexposed cotyledons or fully developed leaves. It is possible
that all potentially dually localized PAPs may hold different
functions in plastid transcription and in photomorphogenesis
as described for PAP5/HMR/pTAC12. It has been surmised
that a subgroup of the PAPs may assemble into a complex
in both the nucleus and the plastid and participate in gene
regulation, by interacting with two different RNA polymerases. Extensive research will be necessary to elucidate these
events in detail. Urgent questions that remain open are those
for the targeting of newly expressed PAPs to nucleus and
plastid, and determining which localization signal becomes
dominant under which condition. Does nuclear localization
require differential translational initiation for removal of the
chloroplastic transit peptide or does trafficking towards the
nucleus occur through a plastidial route in an unknown process, as proposed for Whirly1 (Fig. 1) (Isemer et al., 2012)?
In the PEP complex, one major functional group is comprised of PAPs involved in DNA/RNA metabolism and
gene expression regulation, while the second group is related
to redox regulation and reactive oxygen species protection
(Steiner et al., 2011). The first group includes PAP1, PAP2,
PAP3, and PAP7 (but also PAP5 and PAP12, as described
recently; Kindgren and Strand, 2015). PAP1 is a protein with
a predicted size of 110 kDa and an apparent size of 99 kDa in
SDS-PAGE (Yagi et al., 2012). The gene-derived amino acid
sequence displays a predicted SAP domain that is known to
be involved in DNA/RNA binding (Fig. 3). It also displays
additional domains such as a pentatricopeptide repeat (PPR)
motif and a domain associated with the stabilization of rbcL
transcripts. PPR proteins are known to be involved in RNA
metabolism such as processing, splicing, editing, and translation (Delannoy et al., 2007; Schmitz-Linneweber and Small,
2008; Pfalz et al., 2009). It is reported that pap1 mRNA and
the corresponding protein accumulate within 1–3 h, respectively, after illumination of dark-grown Arabidopsis seedlings
(Yagi et al., 2012). PAP2 is another PPR-containing protein of
89 kDa that displays many structural similarities with PAP1
(Fig. 3) or a small MutS-related mismatch repair domain protein, MRL1, mainly because of the characteristic fold of the
PPR domain (Pfalz et al., 2006; Johnson et al., 2010; Steiner
et al., 2011). A more detailed analysis of the specific functions
of PAP1 or PAP2 regarding their RNA target(s) is, however,
still required. PAP3, a 77 kDa protein with an S1-like domain,
was found to interact with other subunits of the PEP complex
and PAP12 in particular (see below) (Yu et al., 2013). It is
predicted to interact with RNAs through its S1-like domain
(Fig. 3) in Nicotiana benthamiana (Jeon et al., 2012). PAP7/
pTAC14 is a SET domain-containing protein of 49 kDa that
may play a role in the transfer of methyl groups on target proteins (Gao et al., 2011). However, so far there is no evidence
for this kind of function for PAP7/pTAC14.
Other PAPs may fall into the category of subunits necessary for (redox-dependent or redox-related) gene regulation.
PAP6/FLN1 is a protein of 49 kDa and, like its paralogous
protein FLN2, displays a phosphofructokinase domain that
spans the entire length of the protein (Steiner et al., 2011;
Gilkerson et al., 2012). However, a fructokinase activity of
the protein has not been confirmed (Arsova et al., 2010) and
its true function is still elusive. FLN2, interestingly, was not
identified as a PAP and its corresponding knockout mutant
displays a less severe phenotype. It thus represents a more
transient or peripheral component within the TAC (Gilkerson
et al., 2012, Pfalz and Pfannschmidt, 2013). Other protein
modifications are possible within the PEP complex by interaction with the thioredoxin domain of the 12 kDa protein
PAP10/TrxZ, which has been proven to interact with PAP6
(Arsova et al., 2010; Schröter et al., 2010). It might confer
redox regulation of the transcription activity; however, experimental evidence for this is still lacking. Interestingly PAP10
is the first protein for which it was shown that, despite the
specific loss of its functional domain, it still rescues the pap10
mutant phenotype with respect to albinism and plastid gene
expression (Wimmelbacher and Börnke, 2014). This coincides with the proposed model of PAPs as essential structural
components of the PEP complex (see above). Two other paralogous proteins found in the complex are PAP4 and PAP9,
both with iron superoxide dismutase domains (Myouga et al.,
2008). These 26 and 29 kDa proteins may serve as protection
against oxidative stresses generated during the first activities
of the photosynthetic apparatus. A partially functional redundancy is observed in the single mutants, as they show palegreen phenotypes rather than the albino phenotype observed
with the double mutant. Interestingly, PAP4 displays a highly
aberrant apparent molecular weight in SDS-PAGE, suggesting that it might exist as a trimer (Steiner et al., 2011).
Three proteins among the PAPs are less well understood.
PAP11 is a MurE domain-containing protein with a theoretical mass of 85 kDa, as concluded from structure prediction and its degree of homology to bacterial proteins able to
incorporate amino acids into peptidoglycan (Garcia et al.,
2008), but such a function is difficult to reconcile with plastid
transcription. PAP8, a 31 kDa protein, is the most enigmatic
protein of the complex, as it displays no homologies to any
known proteins so far and can only be linked to the formation of the PEP complex by its physical presence within it.
No other data about it are available, but sequence analysis of
the gene identified a potential NLS, suggesting the possibility
of having a second nuclear function. The same is true also
for PAP12, a 12.47 kDa protein that was shown to interact
with PAP3, PAP5, PAP6, and PAP7 in a yeast-two-hybrid
6966 | Pfannschmidt et al.
approach (Yu et al., 2013). Future studies will have to unravel
the precise functions for each of these proteins in the PEP
complex and whether or not the functions are related to
transcription.
Phage-type DNA-dependent RNA polymerases of
plastids (NEPs)
First indications for the existence of a nuclear-encoded plastid-localized RNA polymerase activity came from studies on
plant systems that either do not possess or do not express
the plastid rpo genes due to gene loss or ribosome deficiency
(Morden et al., 1991; Hess et al., 1993). Since plastids in these
plants cannot express the Rpo subunits, they were expected
to be defective in plastid transcription. However, numerous
plastid transcripts were readily detectable, pointing to a second, probably nuclear-encoded enzyme activity. A first candidate for this activity was purified from spinach as a potential
single-subunit RNA polymerase of 110 kDa (Lerbs-Mache,
1993). Further studies using trans-plastomic tobacco lines in
which the rpo genes were genetically inactivated confirmed the
presence of a nuclear-encoded transcription system in plastids that uses a distinct set of promoters (see below). In order
to distinguish between the two activities, the terms nuclearencoded polymerase (NEP) and plastid-encoded polymerase
(PEP) were coined (Hajdukiewicz et al., 1997). Final proof
then came from studies in Arabidopsis that identified two
nuclear genes for a T3/T7 phage-like single-subunit RNA polymerase being targeted to mitochondria and plastids, respectively (Hedtke et al., 1997). These genes were called rpoT for
RNA polymerase of phage T3/T7 type. In Arabidopsis, three
different genes exist that encode RNA polymerases targeted
to mitochondria (RpoTm) and plastids (RpoTp) and, in
addition, according to transit peptide::reporter gene fusion
experiments, an enzyme (RpoTmp) that is targeted to both
organelles (Fig. 1) (Hedtke et al., 2000; Liere et al., 2011).
The gene variants probably emerged from duplication events
multiplying the gene for the mitochondrial RNA polymerase,
redirecting this RNA polymerase activity also to plastids by
acquisition of corresponding transit peptides. However, so
far a gene for RpoTmp has not been detected in monocotyledons, suggesting that it is a late evolutionary acquisition
in dicotyledons (Liere et al., 2011). All rpoT genes encode
phage-type enzymes composed of only a single subunit of
around 110 kDa. In vitro experiments with recombinant
enzyme activities or overexpression of these proteins indicate
faithful promoter recognition, transcription initiation, and
elongation, as well as for a specific transcription initiation factor (Liere et al., 2004; Kühn et al., 2007). Specificity factors
mediating promoter recognition and transcription initiation
as known for yeast or human mitochondria have not been
identified yet in plants (Richter et al., 2010). Nevertheless, the
NEP enzymes specifically recognize three classes of distinct
promoters in vivo designated class Ia, class Ib, and class II
(also called type 1a, type 1b, and type 2). Class Ia contains
a conserved YRT motif several nucleotides upstream of the
transcription start site (Liere and Maliga, 1999). Class Ib contains an additional GAA box around 20 nt upstream of the
YRT motif, and class II represents all promoters without the
YRT motif (Weihe and Börner, 1999). These distinct types of
promoters suggest the existence of protein factors that specifically direct the NEP enzymes to these sites, and at least one
protein factor called CDF2 has been reported in spinach that
is able to direct RpoTmp to the PC promoter of the rRNA
operon (Bligny et al., 2000) (see also below). However, no
specificity factor could be unambiguously identified by mass
spectrometry or other techniques. A cross-interaction with
specificity factors of the PEP enzyme as well as with the PEP
complex itself also appears very unlikely, as NEP enzymes
have never been identified in any mass spectrometry analysis
of PEP complexes. Furthermore, neither NEP nor sigma factors (with the exception of Sig2), or eukaryotic transcription
factors with predicted plastidial localization (Wagner and
Pfannschmidt, 2006), have ever been identified in nucleoidenriched fractions or TAC preparations (Pfalz et al., 2006;
Majeran et al., 2012; Melonek et al., 2012). This lack of evidence suggests that NEP enzymes as well as transcription
regulators are probably present in very low abundance or at
very restricted time points outside the harvesting time point
of current mass spectrometry studies.
The regulation of NEP activity is still an open question.
NEP activity cannot be inferred from the protein level since it
has been shown that during plastid maturation the NEP protein level decreases but its activity increases (Cahoon et al.,
2004). In addition, plastidial RpoTmp activity differs in different plant species. While in Arabidopsis RpoTmp activity
is limited to the seed imbibition stage and declines rapidly
after cold release (Courtois et al., 2007), in spinach it can be
detected during all developmental stages (Bligny et al., 2000;
Azevedo et al., 2008). Some results suggest that the intraplastidial localization of the enzyme might play a role in its activity in spinach. For instance, immunodetection with specific
antisera detected RpoTp and RpoTmp in both stroma and the
membrane fraction of spinach chloroplasts (Azevedo et al.,
2006). These studies demonstrate an increase in thylakoid
membrane association of RpoTmp during leaf development
that is mediated by a RING H2-protein named NIP for NEP
interacting protein. A regulatory role of this membrane association was suggested because it correlates with a decrease in
rDNA transcription (Azevedo et al., 2006, 2008). In contrast,
regulation of RpoTmp activity in Arabidosis should be different. RpoTmp exclusively transcribes the rRNA gene operon
at the PC promoter, and this activity is limited to the period
of seed imbibition (Courtois et al., 2007).
The plastid-localized NEP polymerases are very active at
early steps of seed germination (Demarsy et al., 2006, 2012;
see below), and it is at this specific time that they are intensively expressed. In particular, accumulation of RpoTmp
transcripts is strongly increased during seed stratification and
then declines after root protrusion (Demarsy et al., 2006).
Mitochondrial RpoTmp activity was found to be essential for
the expression of a subset of mitochondrial genes coding for
the subunits of respiratory chain complexes I and IV (Kühn
et al., 2009). These expression data were obtained at an early
step of plant development (when the first pair of leaves comes
out) and at the rosette stage. However, it was not investigated
Plastid RNA polymerases govern chloroplast biogenesis | 6967
whether the mitochondrial RpoTmp enzyme is also active
during seed germination, as shown for the plastid equivalent.
Interestingly, these early steps of seed germination are
characterized by a strong production of anti-sense RNAs
(Demarsy et al., 2012). NEP polymerases are known to
exhibit a high readthrough transcription activity (Legen et al.,
2002). This might cause the generation of anti-sense RNAs
that are complementary to coding regions that are placed on
the opposite DNA strand. As suggested for the psbT gene,
hybridization of anti-sense RNAs with the corresponding
sense RNAs might inhibit translation (Zghidi-Abouzid et al.,
2011). In addition, in plants where the expression of RNase J
was silenced, anti-sense RNAs were accumulated at high levels
(Sharwood et al., 2011). This increased presence of anti-sense
RNAs was causing a strong blockage of translation in these
silenced plants. Interestingly, in situations where the NEP
activity is very high, as for seeds at early steps of germination
(Demarsy et al., 2012) and tobacco rpo-knockout plants that
are deficient in PEP activity (Legen et al., 2002), transcripts
were also not translated. Whether the anti-sense RNAs are
just an unspecific by-product of the NEP readthrough transcription activity or whether they possess a distinct role in
regulation of translation is not yet understood.
Investigation of chloroplast development
in plants
Light-dependent chloroplast development in vascular plants
has been investigated extensively as part of photomorphogenesis and represents a prime example for the influence of
light on developmental programmes in biological systems.
The most common biological test system for chloroplast
development used over the years is the light-induced de-etiolation of dark-grown seedlings when shifted from dark to the
light (Pogson and Albrecht, 2011). Dark-grown angiosperms
such as Arabidopsis and Sinapis typically perform a special
developmental programme called skotomorphogenesis exhibiting hypocotyl elongation, formation of an apical hook, and
yellow cotyledons that contain a dark-specific plastid type,
the etioplast. These plastids lack a thylakoid membrane system but instead possess a so-called prolamellar body, which
is comprised of paracrystalline structures of protochlorophyllide, NADPH, and the enzyme protochlorophyll-oxidoreductase. Upon illumination, chlorophyll biosynthesis
and thylakoid membrane formation are initiated, and within
a few hours a fully developed chloroplast is generated while
in parallel the apical hook is opening and hypocotyl elongation is stopped (Solymosi and Schoefs, 2010). In Arabidopsis,
numerous mutants with defects at different steps of these
developmental programmes have been developed, and essential regulators such as constitutive photomorphogenesis in the
dark (cop) (a repressor of photomorphogenesis) or hy5 (the
major activator of the light-dependent response) were identified (Casal and Yanovsky, 2005). Extensive research led to
an understanding and build-up of the complex regulatory
networks behind these light-dependent developmental programmes (Jiao et al., 2007), and chloroplast biogenesis is
mainly understood as a consequence of the action of these
networks.
The etioplast–chloroplast transition is, however, only one
case of chloroplast biogenesis among others. Under natural
conditions, chloroplast biogenesis in seedlings occurs more
often as a concomitant process of germination without a prolonged dark phase, since the seeds often are not fully covered
from the light. Furthermore, while studies on seedling development are well accepted as developmental models, these
approaches neglect the chloroplast formation in growing or
mature plants. Here, chloroplasts are permanently generated in all meristems of the aerial and illuminated parts of
a plant starting from proplastids in the meristematic stem
cells. Recent studies of chloroplast formation in the shoot
apical meristam of Arabidopsis have revealed very dynamic
and tissue-dependent revertible chloroplast development
(Charuvi et al., 2012). These observations suggest that, in
plants beyond the seedling stage, modified or different regulation pathways of chloroplast biogenesis might be active that
cannot be fully represented by studies on seedlings. It must be
noted that the primary steps of chloroplast biogenesis occur
only in meristematic cells, while in mature green cells chloroplasts multiply by division without running through the early
developmental programme.
Finally, Arabidopsis (but also many other plants) displays
a transient greening stage of the embryo during seed formation within their siliques (Allorent et al., 2013). This includes
chloroplast formation within the early embryo, which is then
reversed by de-differentiation into so-called eoplasts in the
late embryo during the desiccation of the seed. It should
be noted that, while in Arabidopsis the maturing seeds turn
to a brown colour, other plants maintain their seeds in a
green stage until full maturation of seeds, provoking a question about the function of chloroplasts during this stage
(Borisjuk et al., 2013). The primary chloroplast biogenesis in
Arabidopsis embryos is largely not understood and requires
further detailed investigation (see below).
Additional complexity in this context is introduced by
the differences in plastid development between monocotyledonous and dicotyledonous plants. Leaf development in
monocotyledonous plants depends on the activity of a basal
meristem leading to an age-dependent gradient of cells and
plastids from the bottom towards to the tip of the leaf (for a
recent review, see Pogson et al., 2015). In conclusion, we need
to learn much more about the decisive and limiting steps in
chloroplast biogenesis of terrestrial plants. One major aspect
(and the focus of this review) is the interaction of the various
RNA polymerases within the genetic compartments of the
plant cell, which are required for the faithful formation of
chloroplasts.
Regulation and interplay of plastid RNA
polymerases during embryogenesis,
germination, and seedling development
The initial model for the interaction of NEP and PEP enzymes
has been developed based on research data obtained in
6968 | Pfannschmidt et al.
transplastomic tobacco lines in which plastid rpo genes were
inactivated by the insertion of an aadA gene cassette (Allison
et al., 1996; Hajdukiewicz et al., 1997). Homoplastomic tissues and lines of these Δ rpo-plants are deficient in plastid
PEP activity but retain the NEP enzyme activity (see above),
allowing a discrimination between PEP- and NEP-dependent
transcription units. It was proposed that the NEP enzyme
is responsible for the early expression of so-called ‘housekeeping’ genes (including rpo genes and other gene expression-related genes), while the PEP enzyme transcribes genes
required for later chloroplast biogenesis such as the genes for
the photosynthetic complexes (Hajdukiewicz et al., 1997).
In simple terms, the model proposes that the NEP enzyme
transcribes the rpo genes on the plastome of proplastids or
non-green plastids in meristematic cells. The resulting PEP
enzyme then becomes important for the expression of photosynthesis genes and the build-up of the plastid photosynthetic apparatus. This proposed sequence of action of the two
types of RNA polymerases is mainly based on their differential use of different types and/or combinations of promoter
elements (see above). According to this usage, plastome genes
were subdivided into three classes that possess promoters
only for PEP (class I) (photosystem genes), for PEP and NEP
(class II) (most other genes), or only for NEP (class III) (the
rpo operon, accD and ycf2) (see Fig. 1). This model was a
major breakthrough as it could explain the complex process
of chloroplast biogenesis as a simple sequential action of
different RNA polymerases activating different gene groups
depending on the developmental stage.
Data and observations raised in the last 15 years expanded
our knowledge about the action of PEP and NEP transcription systems. There is growing evidence that both polymerases
are active at the same time in all green or non-green tissues
but with different degrees of activity. Systematic comparison
of plastid transcription in green and white tissues of ΔrpoA
tobacco plants and in primary leaves in the barley mutant
albostrians revealed that PEP is the predominant transcriptional activity in green tissues, generating most of the mRNAs,
tRNAs, and rRNAs (Legen et al., 2002; Zhelyazkova et al.,
2012). The NEP enzymes display a minor activity in green
tissues but are enhanced in white tissues of the mutants
where they transcribe even photosynthesis genes from remote
promoters, although to a low degree (Courtois et al., 2007;
Zhelyazkova et al., 2012). An enhanced NEP activity in white
plastids of PEP-deficient mutants appears to be characteristic for this type of mutants, as it has also been reported in
many other albino mutants and especially in mutants with
pap gene expression defects (Pfalz and Pfannschmidt, 2013).
Apparently, PEP deficiency causes a condition in plastids
that leads to either a compensatory or accidental enhancement of NEP activity, although this cannot rescue the chloroplast biogenesis programme. A recent study performed in
maize proposes a predominant role of PEP in the generation of tRNAs supporting active translation in chloroplasts
(Williams-Carrier et al., 2014). This coincides with the low
levels of photosynthesis proteins found in white plastids. At
the same time, this negatively affects the chlorophyll biosynthesis pathway, since the glutamyl-tRNA trnE represents the
precursor of aminolevulinic acid, the entrance substrate of
the tetrapyrrole biosynthesis pathway (Rüdiger and Grimm,
2006; Tanaka and Tanaka, 2007). These and other new data
(e.g. about the requirement of PAP subunits) allow refining
of the initial sequential model of NEP and PEP activity and
adapting it to the timely and spatial dynamics that occur during tissue development in the life cycle of a plant.
Action of RNA polymerases during embryogenesis
The plant life cycle of a diploid dicotyledonous angiosperm such as Arabidopsis starts with the fertilization of the
egg cell, resulting in a zygote. This zygote develops via distinct gene expression programmes and developmental steps
(octant, globular stage, heart stage, and torpedo stage) into
a fully developed plant embryo comprising two cotyledons,
a shoot axis and a primary root embedded into a seed shell
(Le et al., 2010). In Arabidopsis but also in many other plants,
this involves a transient photosynthetic stage in which the
complete embryo turns green and generates fully developed
chloroplasts. The function of these chloroplasts is not fully
understood yet, but repression of this photosynthetic phase by
electron transport inhibitors (e.g. DCMU) negatively affects
the germination ability of the resulting mature seeds (Allorent
et al., 2015). The development of the embryo, still connected
to the mother plant, might be influenced by hormones from
the mother plant at this stage, but may also include independent processes that are controlled by the embryo itself. A clearcut separation of these influences is, however, difficult at
this stage. Based on the observations given above, it must be
assumed that all steps of the RNA polymerase-driven expression cascade for the full development of a chloroplast become
active in the embryo. This includes the activation of NEP and
PEP enzymes, as well as the expression of interacting sigma
factors and PAPs. Indeed, expression and action of both
NEP and PEP activities were identified in this particular stage
(Allorent et al., 2013). Likewise activation of pap gene expression could be identified in publicly available expression data
sets during the embryonic stage of Arabidopsis (Kremnev
and Strand, 2014) (Fig. 2). As already mentioned above,
Arabidopsis seeds lose their chloroplasts during maturation
and turn into brown seeds with chlorophyll-less embryos.
This requires an active block of chloroplast function followed
by degradation of chloroplast membrane structures, protein
components, and pigments within the embryo. Seed maturation, desiccation, and dormancy are largely determined by
hormonal regulation via abscisic acid from the mother plant.
A recent study reports that abscisic acid induces the expression of RSH2 and RSH3, two plastid-localized enzymes
capable of synthesizing guanosine tetraphosphate (ppGpp),
an alarmone that induces the stringent response in bacteria
(Yamburenko et al., 2015). This ancient signalling pathway
of probable endosymbiotic origin has been retained in plants;
however, no real physiological function has been attributed
to plant ppGpp yet (van der Biezen et al., 2000; Takahashi
et al., 2004). As for bacterial RNA polymerases, the molecule
was shown to inhibit transcriptional activities of purified
PEP preparations in vitro (Sato, 2009). It is thus conceivable
Plastid RNA polymerases govern chloroplast biogenesis | 6969
that abscisic acid shuts down PEP activity in chloroplasts of
embryos during seed maturation by enhancing ppGpp production. This would lead to a halt in gene expression and
could initiate subsequent disassembly of the photosynthetic
apparatus. It has been shown recently that the plastids in the
desiccated seeds do not fully return to the proplastid stage
but retain the transcriptional apparatus at both the protein
and transcript level in order to allow an immediate regain of
transcription during germination and subsequent seed development (Allorent et al., 2013).
Action of RNA polymerases during germination and
plant development
Upon imbibition and germination of the seed, a second
phase of chloroplast development can be observed within
the now-growing embryo. This process is, however, restricted
to the tissue of the cotyledons, which is in contrast to the
embryo stage during seed formation where all tissues become
green. Apparently during seedling development, chloroplast
formation in the hypocotyl and root is either actively prevented or not promoted. This would require either expression of an inhibiting factor or the repression of an activating
factor. A recent study suggests that chloroplast formation in
the roots is actively repressed by hormone signals from the
shoot (Kobayashi et al., 2012). This observation also demonstrates that chloroplast formation is a developmental subprogramme that can be switched on and off and does not simply
represent a concomitant process of illumination. In the cotyledons, it has been shown that the same RNA polymerase
expression cascade runs as before in the seed embryo, leading
to a quick generation of chloroplasts in the leaf blade tissues (Demarsy et al., 2006). NEP transcripts both for RpoTp
and RpoTmp are already present in the dry seed (Demarsy
et al., 2006; Allorent et al., 2013). Upon imbibition and germination, NEP transcripts and activities increase and peak at
2–3 d after germination. They trigger a rapid expression of
plastid rpo genes, increasing subsequently PEP activity. This
coincides with a high pap gene expression detected after imbibition of the seed (Fig. 2). Whether NEP and PAP expression is controlled by the same regulators in the nucleus is not
known yet. PEP activity stored in the eoplast after de-differentiation of embryo chloroplasts appears to be important for
efficient germination, as inhibition of this activity by tagetin
treatment delays significantly germination (Demarsy et al.,
2006). However, these starter molecules are apparently not
sufficient to induce full chloroplast development and require
supplementation of activity by generation of novel PEP complexes. This process of complementation probably becomes
interrupted when germination occurs in the dark. In mustard,
this leads to accumulation of the PEP core enzyme, which is
rapidly reconstructed by the addition of PAPs upon illumination (Pfannschmidt and Link, 1994). The precise sequence
of events that occur during this light-triggered reconstruction of PEP is largely not understood and will require careful dissection in future studies. Existing data suggest that
it may include regulation at the levels of transcription,
translation,and even post-translational mechanisms (Steiner
et al., 2011; Yagi et al., 2012). The strong albino phenotype
of pap knockout mutants indicates that the process of PEP
reconstruction is an essential bottleneck in the generation of
chloroplasts. Interruption of any single step in it should lead
to similar phenotypes. A detailed characterization of all pap
mutants as well as additional genetic screens for regulatory
factors will be necessary to fully understand this process. The
high speed at which chloroplasts are generated in dark-grown
seedlings upon illumination suggests that the time window
for the course of events is probably in the hour to minute
range. In the meristems of aerial parts of adult plants, such as
the shoot apical meristem, it should be expected that the processes run even faster, since no artificial pausing due to darkening does occur. A study analysing the presence of thylakoid
membranes in plastids of the different cell layers in the shoot
apex of Arabidopsis could identify fairly developed thylakoids
in the L1 and L3 layers, while in the L2 layer true proplastids
were identified (Charuvi et al., 2012). This suggests that the
differential action of NEP and PEP enzyme must be under a
tight cellular control. The described secondary loss of chloroplasts in epidermal cells of leaves (Charuvi et al., 2012) could
be explained by a targeted repressive effect on PEP activity
e.g. by ppGpp, followed by a specific degradation of plastids similar to the processes described for the late embryonic
phase. How these processes are performed and regulated is
largely unexplored.
The events described above are based mainly on observations in dicotyledonous plants. In principle, it should be
expected that similar processes also occur in monocotyledonous plants, but adapted to the specific developmental gradient of the plastids caused by the action of the basal meristem
(Mullet, 1993). Indeed, many similarities have been reported.
In maize, lack of ZmpTAC2, ZmpTAC10, and ZmpTAC12
(the orthologues of Arabidopsis PAP2, PAP3, and PAP5) each
leads to PEP deficiency exactly as described for Arabidopsis
(Pfalz et al., 2015). Analysis of the barley mutant albostrians
indicates enhanced NEP activities in plastids of tissues with
PEP deficiency (Zhelyazkova et al., 2012). Nevertheless, there
also exist differences from dicotyledonous plants, for instance
the existence of two variants of ZmpTAC12 (Pfalz et al.,
2015). Whether this represents a species-specific difference or
a difference between the plant clades needs to be elucidated
in the future.
Concluding remarks
The transcription of plastidial genes is essential for all plastid types regardless of whether they perform photosynthesis
or not. Proper plastid development therefore involves in all
cases the action of the plastid RNA polymerases. It should
be noted that, in green tissues with mature chloroplasts, the
organelles do multiply by fission rather than by biogenesis
from proplastids. The role of plastid RNA polymerases here
is different from in meristems, embryos, or cotyledons, and is
mainly restricted to the maintenance of transcript levels to
allow sufficient protein production in the mature chloroplast.
Future studies focusing on plastid development will therefore
6970 | Pfannschmidt et al.
need to address a complete understanding of functioning and
regulation of plastid RNA polymerases in both a spatial and
time-resolved manner at the cellular and tissue levels. This
will also include the search for essential (master?) regulators
controlling the expression of the nuclear-encoded components of the plastid transcription machineries (NEPs, Sig1–6,
and PAPs) and putative signalling molecules for retrograde
signals from the plastids. Identification of such regulators
and signalling molecules may help us to understand the coupling (and its limitations) between plastid development and
developmental programmes that control plant morphology
and provides interesting targets for potential biotechnological applications. A detailed understanding of the action of
plastid RNA polymerases will be essential for future ‘green’
biotechnology and bioenergy applications. This includes
functional and structural (crystallographic) analyses of the
complex. Necessary experimental tools are readily available,
and corresponding working models await experimental proof
promising fundamental new discoveries in this research field.
Acknowledgements
Work in Team 7 ‘Nucleo-plastidic interaction: Chloroplast biogenesis and
redox control’ at the Laboratoire de Physiologie Cellulaire & Végétale
(LPCV) was funded by the DFG to TP (Pf323-4 and Pf323-5), by Université
Grenoble-Alpes (AGIR2015) to RB and the Labex GRAL (Grenoble
Alliance for Integrated & Structural Biology) to LPCV. Copyright for colorized scanning electron microscopy images of the Arabidopsis flower used
in Fig. 2: J. Berger and D. Weigel, Max-Planck-Institute for Developmental
Biology, Tübingen, Germany.
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